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Endocrinology Vol. 145, No. 11 4853-4865
Copyright © 2004 by The Endocrine Society

Role for a Novel Signaling Intermediate, Phosphatidylinositol 5-Phosphate, in Insulin-Regulated F-Actin Stress Fiber Breakdown and GLUT4 Translocation

Diego Sbrissa, Ognian C. Ikonomov, Jana Strakova and Assia Shisheva

Department of Physiology, Wayne State University School of Medicine, Detroit, Michigan 48201

Address all correspondence and requests for reprints to: Assia Shisheva, Department of Physiology, Wayne State University School of Medicine, 540 East Canfield, Detroit, Michigan 48201. E-mail: ashishev{at}med.wayne.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The cellular functions and regulation of phosphatidylinositol (PtdIns) 5-phosphate (5-P), the newest addition to the family of phosphoinositides (PIs), are still elusive. Here we have examined a plausible role of PtdIns 5-P as a signaling intermediate in acute insulin action. A wortmannin-insensitive transient increase of PtdIns 5-P mass levels that peaked at 10 min, and declined 20–30 min after insulin stimulation, was observed in both Chinese hamster ovary (CHO)-T cells stably expressing the insulin receptor and 3T3-L1 adipocytes. Similarly to insulin, found to induce a rapid disassembly of Texas-Red phalloidin-labeled actin stress fibers in CHO-T cells, microinjected PtdIns 5-P, but not other PIs, decreased the number and length of F-actin stress fibers in this cell type to a magnitude seen in response to insulin. Likewise, increases of PtdIns 5-P by ectopic expression of the PtdIns 5-P-producing enzyme PIKfyve yielded a similar effect. As with insulin, the PtdIns 5-P-induced loss of actin stress fibers was independent of PI 3-kinase activation. Furthermore, sequestration of functional PtdIns 5-P, either by ectopic expression of 3xPHD domains that bind selectively PtdIns 5-P or by microinjecting the GST-3xPHD fusion peptide, abrogated insulin-induced F-actin stress fiber disassembly in CHO-T cells. In 3T3-L1 adipocytes, microinjected PtdIns 5-P, but not other PIs, partially mimicked insulin’s effect of translocating enhanced green fluorescent protein-GLUT4 to the cell surface. Conversely, insulin-induced myc-GLUT4 vesicle dynamics was arrested in the presence of coexpressed enhanced green fluorescent protein-3xPHD. Involvement of PIKfyve membrane recruitment, but not activation, and/or a decrease in PtdIns 4,5-bisphosphate levels are likely to be among the mechanisms underlying the insulin-induced PtdIns 5-P increase. Together, these results identify PtdIns 5-P as a novel key intermediate for insulin signaling in F-actin remodeling and GLUT4 translocation.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
PHOSPHORYLATED METABOLITES OF phosphatidylinositol (PtdIns), collectively called phosphoinositides (PIs), represent a minor fraction of the eukaryotic cell lipids but play a major regulatory role in diverse cellular processes such as signaling, membrane trafficking, cytoskeletal reorganization, DNA synthesis, and cell cycle (for recent reviews, see Refs. 1, 2, 3, 4, 5, 6, 7, 8). In most cases, PIs serve as regulatory membrane-localized signals to recruit/activate downstream protein effectors that display PI-specific binding domains. A growing number of PI-binding protein modules have been recently identified, including the PH, the FYVE finger (Fab1p, YOTB, Vac1p, and EEA1), the FERM (Four.1-Ezrin-Radixin-Moesin), the Epsin N-Terminal Homology, the PX (Phox Homology) and PHD finger (Plant HomeoDomain) domains (for recent reviews, see Refs. 9, 10, 11, 12, 13). Although their specificity could be broad in some instances, the PI-binding modules have been critical for elucidating the role of PIs in cellular regulation and the mechanism of their function.

Of the seven PIs, the cellular roles of PtdIns 5-phosphate (PtdIns 5-P) are the least well known. This is related to the fact that, in mammalian cells, PtdIns 5-P represents only a minor fraction of PIs and is poorly separated from the abundant PtdIns 4-P upon HPLC resolution. Since its initial discovery as a substrate for type II PI phosphate kinases (PIPKs) (14), PtdIns 5-P has been the subject of several studies addressing its role and regulation of its metabolism in a cellular context. This was made possible, in part, due to the implementation of an alternative approach for PtdIns 5-P intracellular determination, the mass assay, resting upon the substrate preference of type II PIPKs for PtdIns 5-P and the effective detection of PtdIns 4,5-bisphosphate (-P2) by HPLC (15). By mass assay, PtdIns 5-P levels were found to acutely increase in response to thrombin stimulation of platelets (15). Conversely, a robust decrease of PtdIns 5-P was seen upon hypo-osmotic shock in mouse 3T3-L1 fibroblasts and adipocytes, thus implicating PtdIns 5-P as a regulatory intermediate in the osmotic response pathway (16). Recently, a marked increase of PtdIns 5-P levels was shown to accompany Shigella flexneri bacteria invasion due to a potent inositol 4-phosphatase activity of the virulence factor IpgD that converts PtdIns 4,5-P2 to PtdIns 5-P and induces dramatic changes of F-actin remodeling in mammalian cells (17). PtdIns 5-P has also been reported to exist and function in the nucleus (18). The PHD finger of the tumor suppressor ING2, a motif common in many chromatin-regulatory proteins, specifically interacts with PtdIns 5-P and was proposed to function as a nuclear PtdIns 5-P receptor to regulate nuclear responses to DNA damage (19). Clearly, although limited, the available studies imply that PtdIns 5-P is a signaling molecule in its own right and functions in the above or other yet-to-be-identified cellular responses.

Insulin stimulates glucose uptake in fat and muscle by inducing the translocation of an intracellular membrane (IM) pool, containing GLUT4 glucose transporters (GLUT4 vesicles), to the cell surface (for recent reviews, see Refs. 7 , 20 , and 21). This is a complex multistep signaling cascade, which is still not completely understood. Initiated by the activated insulin receptor, this cascade appears to require both PI 3- kinase-dependent and -independent signals for optimal performance (for recent reviews, see Refs. 22, 23, 24). Over the last several years, it became apparent that, like other vesicle trafficking events, the filamentous (F)-actin cytoskeleton plays a major role in insulin-regulated GLUT4 vesicle dynamics (25, 26, 27, 28, 29, 30, 31). In fact, insulin has been shown to be a major trigger of F-actin remodeling in a number of cell types that may or may not express insulin-regulatable GLUT4. A rapid decrease of F-actin stress fibers, followed by a transient increase in membrane ruffling (lamellipodia), has been readily observed in insulin-stimulated Chinese hamster ovary (CHO)-T or HIRc-B cells, both expressing the insulin receptor, and 3T3-L1 fibroblasts (29, 30, 32, 33, 34). In differentiated 3T3-L1 adipocytes, which are large, round, lipid-laden cells that do not contain typical stress fibers, increased membrane ruffling and cortical F-actin formation have been seen in response to insulin (28, 29, 30). Insulin-regulated F-actin cytoskeleton remodeling, whether linked or not with GLUT4 vesicle translocation, is likely mediated by two types of signals: one dependent on PI 3-kinase activity, whereas another appears to be PI 3-kinase independent. Here we have tested whether PtdIns 5-P is involved in the mechanisms used by insulin to signal F-actin remodeling and GLUT4 vesicle translocation. We found acutely elevated PtdIns 5-P mass upon insulin stimulation in both CHO-T and 3T3-L1 adipocytes. Insulin-induced disassembly of F-actin stress fibers in CHO-T cells and cell surface redistribution of GLUT4 were mimicked by microinjected PtdIns 5-P in a wortmannin-independent manner and hindered by sequestering intracellular PtdIns 5-P with PHD finger domains. Thus, PtdIns 5-P emerges as a new key signaling intermediate in insulin action.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell cultures
Mouse 3T3-L1 fibroblasts were differentiated into insulin-sensitive adipocytes as previously described (35). CHO-T cells, stably expressing the human insulin receptor, were maintained in Ham’s F-12 medium, containing 10% fetal bovine serum, 50 U/ml penicillin, and 50 µg/ml streptomycin sulfate as specified elsewhere (35).

Glutathione-S-transferase (GST)-PHD constructs and protein production
To generate a GST-3xPHD domain, we used the clone pEGFP-C2–3xPHD of ING2, a kind gift by Dr. Or Gozani (19). The BglII-SalI fragment of the latter construct was ligated in the compatible cohesive ends of a BamH1-SalI digest of pGEX5X-3 in-frame with GST. Escherichia coli strain XA-90 was used for transformation. Production and purification of the GST fusion proteins on GSH-agarose beads (Sigma, St. Louis, MO) were performed essentially as previously described (36). The concentration and quality of the eluted purified proteins were determined electrophoretically by the intensity of the Coomassie-stained protein bands vs. BSA standard (Pierce, Rockford, IL).

Transient transfection
CHO-T cells were transfected with the pEGFP-3xPHD constructs (kind gift by Or Gozani; Ref. 19) using LipofectAmine as a transfection reagent. Twenty hours post transfection, the cells were serum-starved for 4 h and, after microinjection and/or treatments indicated in the figure legends, were processed for fluorescence microscopy analysis. In some experiments, after transfection, CHO-T cells were serum-starved for 12 h in media supplemented with 0.5% BSA before microinjection and stimulation, yielding similar results. Differentiated 3T3-L1 adipocytes were transfected on d 7 of the differentiation program with the pEGFP-GLUT4 (a kind gift by Jeff Pessin) or pEGFP-3xPHD or cotransfected with pcDNA 3.1–7xMyc-GLUT4 (a kind gift by Kostya Kandror) and pEGFP-3xPHD cDNA constructs using the electroporation method as we detailed previously (35). Experiments were performed 20 h post transfection, subsequent to serum starvation (4 h).

Cell treatment and PtdIns 5-P mass assay
Cells were serum-starved (~4 h) in DMEM or Ham’s F-12 media and then treated at 37 C with or without insulin (100 nM) for the indicated time periods. In some experiments, cells were treated with wortmannin (100 nM; 20 min) before insulin. PtdIns 5-P mass assay was performed as we previously specified in detail (16). Briefly, PIs were isolated from cell lipid extracts on neomycin-coated glass beads. Samples, supplemented with PtdIns as carrier, were subjected to PtdInsP-conversion assay using bacterially produced and purified recombinant His-tagged type IIß PIPK (cDNA was a kind gift by Richard Anderson) in a buffer consisting of 50 mM Tris-HCl (pH 7.4), 80 mM KCl, 10 mM magnesium acetate, 2 mM EGTA, 0.01% sodium deoxycholate, and 5 µM [{gamma}-32P]ATP (5 µCi). In some cases, GST-3XPHD domain or GST peptide fragments (each at 2 µg) were added to the assay. The reaction, continued for 1 h at 30 C, was stopped with 200 µl 1-N HCl and extracted with 160 µl chloroform:methanol (1:1; vol/vol). Lower layers were washed and then spotted on an oxalate-treated and activated thin-layer chromatography (TLC) plate (Whatman, PE SIL G, 250 µm). Plates were developed in 65:35 (vol/vol) n-propanol-2 M acetic acid and exposed with Kodak X-omat (Rochester, NY) autoradiography film. For quantitation, 2–10 pmol PtdIns 5-P standards were processed in parallel. Control samples with no enzyme and/or no lipids were run in each experiment. Because type II PIPK can also convert PtdIns 3-P to some degree, the identity of the PtdIns 4,5-P2 radioactive spots at both basal or insulin-stimulated conditions was confirmed by HPLC-inositol head group analysis after lipid extraction from the silica scrapings and deacylation.

Cell metabolic labeling with [32P]orthophosphate and lipid extraction
Serum-starved (1 h) 3T3-L1 adipocytes or CHO-T cells were washed in phosphate-free DMEM and then labeled for 3 h at 37 C in phosphate-free, serum-free DMEM supplemented with 0.5% BSA, 2 mM sodium pyruvate, and 0.8 mCi/ml of [32P]orthophosphate as described previously (37). Cells were stimulated at 37 C with or without insulin (100 nM) for 10 min, and then washed with ice-cold PBS containing protease (1 mM phenylmethylsulfonylfluoride, 1 mM benzamidine, 5 µg/ml leupeptin, 5 µg/ml aprotinin, and 1 µg/ml pepstatin) and phosphatase inhibitors (50 mM NaF, 10 mM sodium pyrophosphate, 25 mM sodium ß-glycerophosphate, and 2 mM sodium metavanadate) and scraped with CH3OH/1 M HCl (1:1; vol/vol) in the presence of 5 mM EDTA and 5 mM tetrabutylammonium hydrogen sulfate. Extracted radiolabeled lipids were deacylated as previously specified (16, 37) and analyzed by HPLC (see below).

Microinjections, fluorescence, and immunofluorescence microscopy
Microinjection in single CHO-T cells, seeded on glass coverslips in 35-mm dishes (Corning Inc., Corning, NY), was performed as described previously (38, 39). Briefly, PIs [all in dipalmitoyl form from Echelon, except for di-C16-PtdIns 3-P, which was from Matreya and di-C16-PtdIns 3,4,5-trisphosphate (-P3), from Sigma] were sonicated and mixed with goat IgG (2.8 mg/ml; Jackson Immune Research Laboratories, West Grove, PA) to allow detection of injected cells. Purified recombinant protein samples were concentrated in the injection buffer (100 mM KCl, 4 mM sodium phosphate, pH 7.4) to 3 mg/ml protein and mixed with goat IgG to visualize injected cells. Before microinjection of serum-starved cells, the cell culture dish was filled with 5.0 ml prewarmed starvation medium supplemented with 20 mM HEPES (pH 7.4) and placed in a dish heater maintaining media temperature constant (36 C) by a temperature controller (Warner Instruments, New York, NY). The reagents were microinjected into the cytosol within a period of 20 min (~50 microinjected cells). Where indicated, cells were treated with insulin (100 nM; 10 min) and/or pretreated with wortmannin (100 nM; 20 min). After microinjection of the protein samples, cells were returned to the CO2/O2 incubator to recover for approximately 45 min and then treated with insulin. Cells were fixed in 4% formaldehyde and double stained with fluorescein isothiocyanate (FITC) antigoat IgG and rhodamine-phalloidin (Molecular Probes; Eugene, OR) to visualize injected cells and F-actin cytoskeleton, respectively. Coverslips were mounted on slides using the Slow Fade Antifade Kit (Molecular Probes). Fluorescence analysis was performed in a Nikon Eclipse TE 200 inverted fluorescence microscope (Mager Scientific, Dexter, MI) using a 60 x 1.4 oil immersion lens and a standard green fluorescence filter for GFP. Images were captured with a SPOT RT Slider charge-coupled device camera (Diagnostic Instruments, Sterling Heights, MI) mounted on the microscope.

Lipid microinjection in differentiated, pEGFP-GLUT4 electroporated, serum-starved 3T3-L1 adipocytes was performed in a similar manner except for mixing sonicated lipids with Texas-Red dextran (70,000; Molecular Probes) for visualization of microinjected cells, as described previously (38). Individual cells were monitored live within a 30-min postmicroinjection period with a Nikon Eclipse TE 200 inverted fluorescence microscope using a Hoffman Modulation contrast system with a x40 objective and a standard green fluorescence filter for GFP. Cells were subsequently stimulated with insulin (100 nM) for 20 min and imaged live as described above. Images were captured with a SPOT RT Slider charge-coupled device camera. Serum-starved 3T3-L1 adipocytes cotransfected with pcDNA3.1–7xMyc-GLUT4 and either pEGFP-C2–3xPHD or pEGFP-C2–3xPHD3K cDNAs were stimulated with insulin and fixed in 4% formaldehyde as described previously (35). Cells were visualized by the enhanced green fluorescent protein (EGFP) fluorescence and a monoclonal anti-Myc antibody (ATCC, Manassas, VA; CRL 1729) that was subsequently detected with Texas-Red conjugated antimouse IgG. Coverslips were mounted on slides, and the fluorescence analysis was performed in a Nikon Eclipse TE 200 inverted fluorescence microscope as described above.

Evaluation and quantitation of actin stress fibers and GLUT4 translocation in single cells
Consistent with previously published criteria (32, 34), individual CHO-T cells displaying parallel actin fibers that colocalized with the nucleus were scored positive for F-actin stress fibers, whereas those showing actin staining in the periphery were scored negative for F-actin stress fibers but positive for membrane ruffles. Injected or transfected individual 3T3-L1 adipocytes were scored positive for GLUT4 vesicle translocation if appearance of a plasma membrane (PM) rim of GLUT4 fluorescence was documented. F-actin- or cell-surface-GLUT4-positive cells were presented as percentage of the total number of microinjected or transfected cells, given in the figure legends. Cells evaluated for F-actin or GLUT4 PM redistribution were scored independently by two observers.

3T3-L1 adipocyte subcellular fractionation and GLUT4 translocation
After transfection (24 h) with indicated 3xPHD cDNA constructs, 3T3-L1 adipocytes were serum-starved (4 h), treated with insulin (100 nM, 20 min), and then subjected to subcellular fractionation exactly as described in Ref. 35 . Intracellular membrane (IM) and PM fractions were subjected to SDS-PAGE and Western blotting with anti-GLUT4 polyclonal antibodies (a kind gift by Mike Czech) as described previously (35).

HPLC analysis and data quantitation
Deacylated 32P-labeled lipids were analyzed by HPLC on a Whatman 235-mm x 4.60-mm column packed with 5-micron Partisphere SAX (H2PO4–) and eluted with a shallow ammonium phosphate gradient at a flow rate of 1.0 ml/min as detailed elsewhere (16, 37). Coinjected internal HPLC standards were prepared or purchased from sources detailed previously (16, 37). Fractions were collected every 0.25 min, and their radioactivity was analyzed simultaneously for 3H- and 32P-labeled standard and products, respectively, with 2.0 ml of ScintiVerse liquid scintillation cocktail on a liquid scintillation counter (Packard Instrument Co., Inc.). The radioactivity of the TLC-scraped [32P]glycerophosphorylinositol (GroPIns) 4,5-P2 formed during PtdIns 5-P conversion assay was analyzed with an on-line flow scintillation analyzer (Packard, Meriden, CT; Radiomatic 525TR).

The radioactivity of the PI peaks was quantified by area integration and is presented as a percentage of the radioactivity determined in corresponding control samples or as a percentage of total PI radioactivity, as indicated in figure legends. Because the separation of PtdIns 5-P from PtdIns 4-P was not complete, for quantitation of the counts under the PtdIns 5-P peak riding on top of the PtdIns 4-P tail, the peak was skimmed and presented as percent of the radioactivity determined in control samples by the same approach.

PIKfyve lipid kinase activity
The in vitro activity was analyzed using anti-PIKfyve immunoprecipitates, derived from lysates of insulin-treated (100 nM, 10 min, 37 C) or control cells, which were incubated with PtdIns and [{gamma}-32P]ATP for 15 min at 37 C. Radiolabeled products were extracted and analyzed by TLC as previously described (38, 40).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Insulin transiently increases PtdIns 5-P mass levels
To examine a plausible effect of insulin on the dynamics of PtdIns 5-P intracellular production, we took advantage of the mass assay that allows one to quantify PtdIns 5-P mass levels in cells (16). We used two types of insulin-sensitive cells: 3T3-L1 adipocytes, considered as prototypic insulin-responsive cells; and CHO-T cells stably expressing the human insulin receptor. As illustrated in Fig. 1Go, both cell types demonstrated marked increases of PtdIns 5-P mass levels upon acute insulin treatment, as judged by the amounts of PtdIns 4,5-P2 synthesized in vitro from extracted PtdIns 5-P and the action of type II PIPK. The effect reached a maximum after 10 min of insulin stimulation, exceeding the basal PtdIns 5-P mass levels by 2.5-fold and 4.2-fold in 3T3-L1 adipocytes and CHO-T cells, respectively (Fig. 1Go, C and D). The effect was transient in both cell types and returned toward the basal levels after 20–30 min of insulin challenge (Fig. 1Go, A–D). Importantly, CHO-T cell pretreatment with wortmannin did not abolish the insulin-induced PtdIns 5-P elevation (Fig. 1EGo). Calculation of the PtdIns 5-P mass levels showed substantially higher amounts (~70-fold) of basal PtdIns 5-P in 3T3-L1 adipocytes compared with CHO-T cells, equal to 580 ± 70 and 8.5 ± 3 pmol/mg protein in 3T3-L1 adipocytes and CHO-T cells, respectively. Together, these results clearly demonstrate a transient elevation of PtdIns 5-P mass associated with acute insulin stimulation in insulin-sensitive cell types. Importantly, this effect could be detected at very low basal PtdIns 5-P mass (see below).



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  		FIG. 1. Acute insulin induces robust increases of PtdIns 5-P mass levels in 3T3-L1 adipocytes and CHO-T cells in a wortmannin-independent manner. Serum-starved 3T3-L1 adipocytes (A and C) or CHO-T cells (B, D, and E) were treated with or without insulin (100 nM) at 37 C for the indicated time intervals (A–D) or for 10 min subsequent to pretreatment with or without wortmannin (100 nM) for 20 min as indicated (E). Cells were then washed and the lipids extracted as detailed in Materials and Methods. PIs were isolated on neomycin-coated glass beads and subjected to in vitro conversion by type II PIP kinase for 1 h at 30 C in the presence of [{gamma}-32P]ATP. 32P-labeled products were separated by TLC and visualized by autoradiography. Shown are representative autoradiograms (A, B, and E) and quantitation from five (C) and four (D) independent experiments for 3T3-L1 adipocytes and CHO-T cells, respectively, presented as a percentage of PtdIns 4,5-P2 from the corresponding nontreated cells for each individual time point (mean ± SEM). PtdIns 5-P intracellular mass (see text) was calculated from PtdIns 5-P standards run in parallel in each experiment (lane 7 in B, 10 pmol).

 
The dramatic difference in the PtdIns 5-P intracellular levels in 3T3-L1 adipocytes vs. CHO-T cells observed here by mass assay is consistent with our previously published data, reproduced herein (Refs. 16 and 37 , and data not shown) using HPLC inositol-head group analysis for determining the basal 32P-PtdIns 5-P accumulation in [32P]orthophosphate-labeled cells. This analysis found radiolabeled PtdIns 5-P in quiescent 3T3-L1 adipocytes at quite substantial amounts, comprising as much as 12% of PtdIns 4-P, in contrast to undetectable levels in CHO-T cells (16, 37). To determine whether insulin-dependent increases of PtdIns 5-P levels could be detected by similar analysis, deacylated lipids, extracted from serum-starved 32P-labeled 3T3-L1 adipocytes or CHO-T cells, stimulated or not, for 10 min with the hormone, were resolved on HPLC columns by a shallow ammonium phosphate gradient, shown previously to give a better resolution of PtdIns 5-P from the descending edge of the PtdIns 4-P peak (16, 40). Using a quantitation method described in Materials and Methods, we detected only small increases of [32P]PtdIns 5-P accumulation in response to insulin stimulation of 3T3-L1 adipocytes and no 32P-PtdIns 5-P accumulation in insulin-stimulated CHO-T cells (Fig. 2Go). This is most likely due to the low basal levels of radiolabeled PtdIns 5-P in the latter cell type that are below the sensitivity of the HPLC-inositol head group detection, as we have demonstrated previously (37). Comparative analysis of data from both HPLC and mass assays reveals that, even at a 4-fold increase, 32P-PtdIns 5-P will represent only approximately 0.75% of the PtdIns 4-P peak in CHO-T cells and will most likely remain buried within the trailing edge of the abundant PtdIns 4-P peak long before the latter reaches base line. Combined data from 3T3-L1 adipocytes and CHO-T cells indicate that HPLC head group analyses are insufficient for measuring basal or stimulated PtdIns 5-P in these and likely in other cell types.



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  		FIG. 2. HPLC analysis reveals 32P-PtdIns 5-P accumulation in response to acute insulin in 3T3-L1 adipocytes but not in CHO-T cells. 3T3-L1 adipocytes or CHO-T cells were serum/phosphate starved for 1 h and then labeled with [32P]orthophosphate for 3 h in phosphate/serum-free DMEM as described in Materials and Methods. Cells were then treated with insulin (100 nM) for 10 min at 37 C or left untreated. Cell lipids were extracted, deacylated, and coinjected with [3H]GroPIns 5-P, [3H]GroPIns 4-P, [3H]GroPIns 3-P, and [3H]GroPIns 4,5-P2 as internal HPLC standards. Fractions, collected every 0.25 min were monitored for [3H] and [32P] radioactivity by liquid scintillation counting. 32P-radioactivity was plotted and the counts within the elution times corresponding to GroPIns 5-P determined and normalized to the GroPIns 4,5-P2 counts. Shown is a quantitation of HPLC elution profiles with respect to insulin-stimulated [32P]PtdIns 5-P accumulation expressed as a percentage of basal [32P]PtdIns 5-P in 3T3-L1 adipocytes of two independent experiments with similar results. A [32P]PtdIns 5-P peak was not detected in basal or insulin-stimulated CHO-T cells in four independent cell labelings.

 
PtdIns 5-P, but not other PIs, mimics insulin in stress fiber disassembly in CHO-T cells
The observed insulin-induced robust increase of PtdIns 5-P mass, demonstrated above, poses a question about the nature of insulin-regulated cellular processes where PtdIns 5-P operates. One well-established acute insulin response in CHO-T cells is the marked reduction of the number and length of F-actin stress fibers that are seen in serum-starved cells (30, 32, 33). Intriguingly, similarly to insulin-induced changes in PtdIns 5-P mass (Fig. 1Go), the insulin-regulated disassembly of F-actin stress fibers has been found to be transient with a maximum at approximately 15 min (Fig. 3AGo) and restoration of the actin stress fiber network after 30 min of insulin presence (Ref. 32 and this study, not shown). Remarkably, unlike the membrane ruffling that operates in a PI 3-kinase-dependent manner, insulin-induced loss of F-actin stress fibers is shown to proceed by a PI 3-kinase-independent (29, 30) or PtdIns 3,4,5-P3-independent mechanism (41, 42). Therefore, we analyzed the role of PtdIns 5-P in insulin-regulated actin stress fiber disassembly in CHO-T cells. We used several approaches to modulate intracellular PtdIns 5-P levels. First, to increase PtdIns 5-P intracellular levels, the lipid was microinjected in the cytoplasm of serum-starved CHO-T cells. Cells were then fixed, and the organization of F-actin was examined by rhodamine-conjugated phalloidin and fluorescence microscopy. Remarkably, PtdIns 5-P, like insulin, was found to induce stress fiber breakdown (Fig. 3BGo). Quantitation from three independent experiments demonstrated that the potency of microinjected PtdIns 5-P to induce stress fiber breakdown is equal to that of insulin (Fig. 3CGo). To test the specificity of this PtdIns 5-P effect we have examined other PI derivatives under similar conditions. Intriguingly, none of the other PIs tested here was found to display the ability to induce stress fiber breakdown (Fig. 3CGo). In all cases the F-actin stress fiber network displayed a similar appearance to that in the control noninjected quiescent cells (Fig. 3BGo). Furthermore, CHO-T cell pretreatment with the PI 3- kinase inhibitor wortmannin failed to significantly affect the loss of stress fibers not only due to insulin (Ref. 30 and this study, not shown) but also those due to microinjected PtdIns 5-P (Fig. 3BGo). These data are consistent with the notion that, in CHO-T cells, PtdIns 5-P mediates insulin’s effect on stress fiber depolymerization and that this proceeds independently of PI 3-kinase activation. Under the experimental conditions used here, we were unable to observe PtdIns 5-P-dependent membrane ruffling in different optical planes of the cells, indicating differential mechanisms underlying the stress fiber breakdown and membrane ruffling.



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  		FIG. 3. Microinjected PtdIns 5-P, but not other PIs, mimics insulin in stimulating stress fiber breakdown in CHO-T cells. A, Serum-starved CHO-T cells expressing the insulin receptor were treated with insulin (100 nM, 10 min, 37 C) or left untreated. Cells were washed, fixed in formaldehyde, and permeabilized. F-actin was visualized with rhodamine-phalloidin. Note the loss of actin stress fibers upon insulin stimulation. B, Serum-starved CHO-T cells were comicroinjected within a 20-min period with indicated PIs and goat IgG to visualize injected cells. Cells in panels e and f were pretreated with wortmannin (Wort) (100 nM) for 20 min at 37 C and then microinjected. Cells were washed, fixed, permeabilized, and stained for F-actin and goat IgG with rhodamine-phalloidin (a, c, and e), and FITC-conjugated donkey antigoat IgG (b, d, and f), respectively. Individual cells, positive for lipid microinjection were scored for the presence of parallel stress fibers with the cell nucleus as described in Materials and Methods. Shown are representative images from three independent experiments with similar results. Arrows in a, c, and e depict microinjected cells, visualized by FITC-conjugated donkey antigoat IgG (b, d, and f). Note that only the PtdIns 5-P-microinjected cells display loss of actin stress fibers, which is seen in both the absence (panel c, the cell pointed to by an arrow) or presence of wortmannin pretreatment (panel e, the two cells pointed to by arrows). C, Quantitation of positive cells for actin stress fibers after microinjection of indicated PIs, presented as a percentage of the total number of microinjected cells (50–100 cells/condition). Control (con) or insulin (ins)-treated cells were microinjected only with vehicle (goat IgG in injection buffer). Each bar represents the mean ± SEM from three independent experiments.

 
We sought to confirm the observed effect of high PtdIns 5-P levels on the F-actin stress fiber loss by an alternative approach. We have recently demonstrated that the PIKfyve enzyme produces intracellularly PtdIns 5-P, because a HEK293 stable cell line expressing PIKfyveWT at a level 7-fold higher than the endogenous protein displayed approximately 2-fold elevated PtdIns 5-P mass (16). To examine whether increased PtdIns 5-P due to PIKfyve action is also associated with actin stress fiber breakdown, we ectopically expressed PIKfyveWT in CHO-T cells and assessed the F-actin structures by rhodamine-phalloidin labeling of serum-starved cells. We found that expression of EGFP-PIKfyveWT also induced loss of actin stress fibers (Fig. 4Go), whereas control EGFP expression did not (see Fig. 6AGo, a and b). Loss of actin stress fibers was observed in approximately 45% of EGFP-PIKfyveWT-transected CHO-T cells, representing approximately 60% of insulin’s effect on stress fiber disassembly in this cell type (see Fig. 3CGo). Clearly, the combined data indicate that PtdIns 5-P, elevated either by direct lipid microinjection or enzymatically, mimics insulin action on stress fiber breakdown.



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  		FIG. 4. Expression of PIKfyveWT results in stress fiber breakdown in CHO-T cells. Twenty-four hours post transfection with pEGFP-PIKfyveWT and subsequent to serum deprivation, CHO-T cells, stably expressing the insulin receptor, were fixed in formaldehyde, permeabilized, and stained with rhodamine-phalloidin (A), as described in Materials and Methods. PIKfyveWT-expressing cells, determined by the EGFP fluorescence signals (B), were scored for the presence of parallel actin stress fibers with the cell nucleus as described in Materials and Methods. Note the loss of actin stress fibers in the PIKfyveWT-expressing cells (arrowheads), seen in 45% of the transfected cells (insulin-induced stress fiber breakdown occurred in 75% of the nontransfected cells as illustrated in Fig. 3CGo).

 


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  		FIG. 6. 3xPHD peptides inhibit insulin-regulated loss of F-actin stress fiber disassembly. A, CHO-T cells expressing the insulin receptor were transfected with pEGFP alone, pEGFP-ING2–3xPHD, pEGFP-ING2–3xPHD3K, or pEGFP-ACF-3xPHD as indicated. Twenty hours post transfections, serum-starved cells were stimulated with insulin (100 nM) for 10 min at 37 C or left untreated as indicated. Cells were washed, fixed in formaldehyde, and stained with rhodamine-phalloidin to visualize F-actin as described in Materials and Methods. Arrowheads in a, c, e, g, i, and k depict the transfected cells seen in b, d, f, h, j, and l by the GFP fluorescence signals. Note in g and k the transfected cells that show stress fibers despite the insulin treatment. The second arrow in e points to a low-level expressing cell. B, Quantitation of transfected cells positive for actin stress fibers presented as a percentage of the total number of cells transfected with each construct (counted 200 cells/experiment/condition). Each bar represents the mean + SEM from three experiments. C, Serum-starved CHO-T cells were microinjected with GST-ING2–3xPHD peptide (3 mg/ml) mixed with goat IgG to visualize the injected cells. Approximately 45 min after microinjection, cells were stimulated with insulin (100 nM) for 10 min at 37 C or left untreated as indicated. Cells were washed, fixed, and stained with rhodamine-conjugated phalloidin and FITC-conjugated donkey antigoat IgG to visualize F-actin (a and c) and microinjected cells (b and d). Note in c the three noninjected cells at the image top demonstrating the typical loss of actin stress fibers, in contrast to the six injected cells showing multiple F-actin stress fibers despite the insulin treatment.

 
Scavenging intracellular PtdIns 5-P inhibits insulin-regulated actin stress fiber disassembly
If high PtdIns 5-P mimics insulin, then its elimination from cells would be expected to oppose insulin effects. To test this hypothesis, we explored recently identified PHD domain modules, shown to preferentially bind PtdIns 5-P (19). The PHD domain of ING2, a candidate tumor suppressor protein, binds PtdIns 5-P with high affinity but could also interact with PtdIns 4-P and PtdIns 3-P, although to a lesser extent (19). In contrast, the PHD domain of ACF protein is exclusive for PtdIns 5-P, but the binding affinity is lower than that of ING2-PHD (19). Therefore, we tested the PHD domain fingers from both proteins and used their three tandem repeat forms to increase binding (19). We first confirmed, by the PtdIns 5-P mass assay, the ability of recombinantly produced GST-3xPHD domain fusion-peptide fragment to knock down functional PtdIns 5-P. Results presented in Fig. 5Go, demonstrating a dramatic inhibition of in vitro PtdIns 4,5-P2 production from both basal or insulin-stimulated PtdIns 5-P intracellular pools in the presence of GST-ING2–3xPHD purified peptide, but not GST alone, are consistent with the notion that the 3xPHD peptide acts as a powerful PtdIns 5-P scavenger.



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  		FIG. 5. 3xPHD domain sequesters functional PtdIns 5-P. Serum-starved 3T3-L1 adipocytes were treated with or without insulin (100 nM) for 10 min at 37 C. Cells were then washed and the lipids extracted. PIs were isolated on neomycin-coated glass beads, preincubated with GST or GST-ING2–3xPHD peptides (2 µg each) for 10 min, and subjected to in vitro conversion by type II PIP kinase for 1 h at 30 C in the presence of [{gamma}-32P]ATP. 32P-labeled products were separated by TLC and visualized by autoradiography. Shown is a representative autoradiogram of three independent experiments with similar results. Note the marked inhibition of PtdIns 4,5-P2 synthesis in the presence of the GST-ING2–3xPHD peptide (lanes 2 and 4).

 
The ability of 3xPHD peptides to arrest stress fiber breakdown induced by insulin was examined in CHO-T cells ectopically expressing ING2–3xPHD or ACF-3xPHD constructs fused with EGFP. Remarkably, expression of both EGFP-ING2–3xPHD and EGFP-ACF-3xPHD peptides, but not EGFP alone, profoundly blocked the loss of F-actin stress fibers in response to insulin (Fig. 6Go, A and B). The peptides had no significant effect on the stress fiber F-actin structures in quiescent cells (Fig. 6Go, A and B). By contrast, expression of a PtdIns 5-P-binding-deficient version of the EGFP-ING2–3xPHD domain, bearing substitutions in K49, K51, and K56 (3xPHD3K; Ref. 19), had practically no effect on insulin-induced actin stress fiber breakdown in CHO-T cells (Fig. 6Go, A and B).

To confirm the inhibitory effect of PtdIns 5-P scavenging by an alternative approach, we have inspected the organization of F-actin stress fibers in CHO-T cells that were microinjected with the GST-ING2–3xPHD peptide before stimulation with insulin. As illustrated in Fig. 6CGo, whereas the GST-3xPHD domain did not significantly change the actin-stress fiber network in quiescent cells, it almost completely prevented the stress fiber breakdown due to insulin. By contrast, control microinjection of GST alone failed to abort insulin-induced loss of actin stress fibers (data not shown).

Effect of PtdIns 5-P and other PIs on GLUT4 translocation in 3T3-L1 adipocytes
Until recently, PI 3-kinase activation and 3'-polyphosphoinositide production were considered to be necessary and sufficient for insulin to stimulate GLUT4 vesicle translocation from intracellular storage sites to the PM and the subsequent glucose uptake into fat and muscle cells. The observation that cell permeable derivatives of PtdIns 3,4,5-P3 failed to mimic insulin’s effect on glucose transport (43) not only suggested an additional PI 3-kinase-independent signaling pathway in adipocytes (22, 23) but also has proven critical the need for direct lipid-derived studies. Because PtdIns 5-P not only constitutes a substantial subfraction of total PtdInsP in resting 3T3-L1 adipocytes (16) but was also found up-regulated in response to acute insulin stimulation of this cell type (Fig. 1Go), we examined whether microinjected PtdIns 5-P could affect GLUT4 vesicle dynamics. We followed the dynamics of transiently expressed EGFP-GLUT4 shown previously to effectively reflect the behavior of endogenous GLUT4 in these cells (35, 44, 45). In addition to PtdIns 5-P, in this setting we have examined PtdIns 4-P, PtdIns 3-P, PtdIns 3,5-P2, and PtdIns 3,4,5-P3 to test the specificity of the effect. In all cases, 24 h post transfection of 3T3-L1 adipocytes with pEGFP-GLUT4 construct, lipids mixed with Texas-Red dextran were microinjected in the cytoplasm of serum-starved cells. EGFP-GLUT4 vesicle appearance at the cell surface was monitored in single cells over a time course of 30 min, while keeping the dish at 36 C by a temperature controller. At the end of the observation period, cells were stimulated with insulin for an additional 20 min to examine the responsiveness of the individual cells to translocate EGFP-GLUT4 to the cell surface. This last step is essential to avoid an artificial underestimation of the microinjected PIs’ effect due to two reasons. First, even if noninjected, only half of the 3T3-L1 adipocytes display the ability to translocate perinuclear EGFP-GLUT4 to the cell surface in response to insulin, as observed previously in several studies, including our own (35). Second, unresponsiveness due to cell damage associated with the PI microinjection is also possible. Therefore, in the quantitative data, summarized in Fig. 7BGo, we score the appearance of the characteristic PM rim of the EGFP-GLUT4 fluorescence signals due to microinjected PI as well as the individual responsiveness of injected cells to a subsequent insulin challenge. Importantly, under the specified experimental conditions, the only one of the here-examined PIs that displayed an ability to mimic insulin in translocating EGFP-GLUT4 to the cell surface was PtdIns 5-P (Fig. 7Go, A and B). This effect was apparent in approximately 50% of the injected cells that responded positively to a subsequent insulin challenge (Fig. 7BGo). Figure 7AGo (d–f) illustrates one typical cell, in which the cell surface EGFP-GLUT4 appearance is readily seen 30 min after PtdIns 5-P microinjection. The effect is comparable with that observed in a vehicle-microinjected cell after 20 min of insulin challenge (Fig. 7AGo, a–c). By contrast, cells microinjected with PtdIns 3,4,5-P3 did not exhibit a noticeable effect in EGFP-GLUT4 cell surface accumulation under similar conditions (Figs. 7AGo, g–i, and B). Given the GLUT4-PM rim is apparent only 15–20 min after PtdIns 5-P microinjection, together with the lack of visible cell surface EGFP-GLUT4 before microinjection, and kinetic data for slow constitutive exocytosis of GLUT4 in this cell type (~230 min; Ref. 46), these results are consistent with the notion that PtdIns 5-P acts by triggering GLUT4 exocytosis rather than inhibiting GLUT4 internalization. Clearly, these results demonstrate that direct administration of PtdIns 5-P mimics, at least in part, the insulin effect on cell-surface translocation of GLUT4 vesicles.



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  		FIG. 7. PtdIns 5-P microinjection, but not other PIs, mimics insulin-induced GLUT4 translocation in 3T3-L1 adipocytes. A, 3T3-L1 adipocytes were electroporated with pEGFP-GLUT4 as described in Materials and Methods. On the next day, cells were serum-starved, and the EGFP-GLUT4-expressing cells (a, d, and g) were injected with Texas-Red (TxR) dextran in injection buffer alone (b) or mixed with the indicated PIs (e and h). Twenty to 30 min post injection, individual cells were monitored live for EGFP-GLUT4 translocation (d and g). Cells were treated with insulin (100 nM) for an additional 20 min (a–c). Shown are representative images depicting the characteristic GLUT4 PM rim by insulin in a vehicle-injected cell (a–c) and the effect of PtdIns 5-P microinjection (5-P, d–f) or PtdIns 3,4,5-P3 microinjection (3,4,5-P3, g–i) on EGFP-GLUT4 localization. Note that PtdIns 5-P, but not PtdIns 3,4,5-P3 produces a EGFP-GLUT4-plasma-membrane rim in the absence of insulin. B, Quantitation of cells positive for EGFP-GLUT4 cell-surface fluorescence in response to microinjection of TxR alone or TxR mixed with indicated PIs, without (black bars) or with (open bars) a subsequent insulin treatment, expressed as a percentage of the total number of monitored microinjected cells (24 cells for TxR dextran; 14 for PtdIns 3-P; 12 for PtdIns 4-P; 33 for PtdIns 5-P; 14 for PtdIns 3,5-P2, and 22 for PtdIns 3,4,5-P3). Note that only PtdIns 5-P was able to trigger the EGFP-GLUT4 redistribution to the PM, and that only around 30% of the microinjected cells were insulin responsive.

 
The role of PtdIns 5-P in insulin-regulated GLUT4 translocation in 3T3-L1 adipocytes was further substantiated by the ability of the 3xPHD domain of ING2 or ACF to inhibit this effect (Fig. 8Go). As illustrated in Fig. 8Go, A and B, in the presence of expressed EGFP-3xPHD domains, acute insulin was unable to induce cell surface redistribution of ectopically expressed Myc-GLUT4. Conversely, expression of a PtdIns 5-P-binding-deficient version of the EGFP-ING2–3xPHD domain, bearing substitutions in K49, K51, and K56 (3xPHD3K; Ref. 19), had practically no effect on insulin-induced cell surface appearance of myc-GLUT4 (Fig. 8Go, A and B). Similar results were obtained by assessing the endogenous GLUT4 distribution in subcellular fractions derived from 3T3-L1 adipocytes transfected with the PHD domain reagents. Whereas expression of EGFP-ING2–3xPHD3K (Fig. 8CGo) or EGFP alone (not shown) did not affect insulin-induced GLUT4 translocation to the PM fraction, expression of EGFP-ING2–3xPHD was inhibitory to a considerable extent (~40%). The transfection efficiency of these constructs was at approximately 50%, implying that EGFP-ING2–3xPHD expression arrested almost completely insulin’s effect on GLUT4 translocation. Clearly, these results demonstrate that insulin’s effect on PM translocation of GLUT4 is specifically influenced by PtdIns 5-P-binding reagents, consistent with the requirement for PtdIns 5-P.



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  		FIG. 8. 3xPHD peptide, but not PtdIns 5-P-binding-deficient 3xPHD3K mutant, inhibits insulin-regulated GLUT4 vesicle translocation in 3T3-L1 adipocytes. A, 3T3-L1 adipocytes were electroporated to coexpress Myc-GLUT4 with either EGFP alone, EGFP-ING2-3xPHD, or EGFP-ING2-3xPHD3K mutant as described in Materials and Methods. On the next day, cells were serum-starved, then stimulated with insulin (100 nM, 20 min) or left untreated, washed, fixed, and processed for fluorescence microscopy as detailed in Materials and Methods. Expressed GLUT4 was detected with anti-Myc monoclonal antibody and Texas-Red-conjugated secondary antibodies (a, d, g, and j). The expression of GFP-based constructs was visualized by the GFP fluorescence (b, e, h, and k). The images shown are representative from three independent experiments in cells coexpressing the plasmids. Bar, 20 µm. B, Quantitation of cells positive for Myc-GLUT4-cell-surface fluorescence in basal or insulin-stimulated 3T3-L1 adipocytes doubly transfected with pcDNA3.1-Myc-GLUT4 and pEGFP vector alone, pEGFP-ING2-3xPHD, or pEGFP-ING2-3xPHD3K, presented as a percentage of the total number of coexpressing cells for each condition. Results shown are from three independent experiments, with observation of approximately 200 cells/condition/experiment, mean ± SEM. C, 3T3-L1 adipocytes were electroporated to express EGFP-ING2–3xPHD or EGFP-ING2–3xPHD3K as described in Materials and Methods. On the next day, cells were serum-starved, stimulated with insulin (100 nM, 20 min) or left untreated, and then fractionated to obtain IM, PM, and cytosol. Indicated fractions were resolved by SDS-PAGE, and insulin-induced GLUT4 translocation was determined by immunoblotting [Western blot (WB)] with polyclonal anti-GLUT4 antibodies. Shown is a chemiluminescence detection of a representative blot of two independent experiments with similar results. Note the inhibition (30–40%) of insulin’s effect on GLUT4 in both IM and PM under 3xPHD vs. 3xPHD3K expression.

 
PtdIns 4,5-P2 levels and PIKfyve enzymatic activity in insulin-stimulated 3T3-L1 adipocytes and CHO-T cells
Mobilization of several pathways may underlie the insulin- induced robust increase of PtdIns 5-P mass in 3T3-L1 adipocytes and CHO-T cells, demonstrated above. These involve up-regulation of the pathways for 5'-polyphosphoinositide breakdown or PtdIns 5-P synthesis from PtdIns and/or down-regulation of pathways responsible for PtdIns 5-P clearance. We first tested whether the PtdIns 4,5-P2 breakdown could account for the observed insulin-dependent increase of PtdIns 5-P mass. Such a mechanism has been recently reported to underlie the PtdIns 5-P rise in mammalian cells due to S. flexneri invasion (17). We examined the amounts of accumulated radiolabeled PtdIns 4,5-P2 after a 10-min insulin stimulation of serum-starved 32P-labeled 3T3-L1 adipocytes and CHO-T cells. As illustrated in Fig. 9Go, the HPLC-head group analysis of extracted and deacylated radiolabeled lipids failed to demonstrate significant insulin-dependent changes in [32P]PtdIns 4,5-P2 accumulation in both cell types. This result is therefore consistent with a lack of significant insulin-dependent increases of the PtdIns 4,5-P2 breakdown and/or inhibition of the PtdIns 5-P clearance through the type II PIPK pathway. However, it should be emphasized that, due to the markedly greater levels of PtdIns 4,5-P2 vs. PtdIns 5-P (Refs. 16 and 37 and this study), even an insignificant decrease in PtdIns 4,5-P2 could yield substantial increases of PtdIns 5-P. Therefore, subtle, spatially restricted variations in PtdIns 4,5-P2 breakdown by a yet-to-be-identified 4P-phosphatase and/or in PtdIns 4,5-P2 synthesis by the type II PIPK pathway to explain the insulin-induced rise in PtdIns 5-P, could not be ruled out, even in the absence of significant changes in the overall 32P-PtdIns 4,5-P2 accumulation in response to insulin.



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  		FIG. 9. 32P-PtdIns 4,5-P2 intracellular accumulation remains unchanged by acute insulin. 3T3-L1 adipocytes or CHO-T cells were serum/phosphate starved for 1 h and then labeled with [32P]orthophosphate for 3 h in phosphate/serum-free DMEM as described in Materials and Methods. Cells were then treated with insulin (100 nM) for 10 min at 37 C or left untreated. Cell lipids were extracted, deacylated and coinjected with [3H]GroPIns 5-P, [3H]GroPIns 4-P, [3H]GroPIns 3-P and [3H]GroPIns 4,5-P2 as internal HPLC standards. Fractions, collected every 0.25 min, were monitored for [3H] and [32P] radioactivity by liquid scintillation counting. 32P-radioactivity was plotted and the counts within the elution times corresponding to the [32P]GroPIns peaks determined by the above [3H]-labeled standards were summed (total radioactivity). Shown is a quantitation of HPLC elution profiles with respect to 32P-PtdIns 4,5-P2, presented as a percentage (mean ± SEM) of total radioactivity of three (for 3T3-L1 adipocytes) or two independent experiments (CHO-T) with similar results.

 
In a cellular context, PIKfyve is the enzyme responsible for the synthesis of PtdIns 5-P from PtdIns (16). Possible insulin-regulated PIKfyve activation in the time course of the observed elevated PtdIns 5-P production in response to insulin was tested in both 3T3-L1 adipocytes and CHO-T cells. In agreement with our previous observation in insulin-stimulated 3T3-L1 adipocytes, insulin action in CHO-T cells did not result in significant changes of the in vitro synthesized PtdIns 5-P by PIKfyve immunoprecipitates derived from cell lysates of basal or stimulated cells (Ref. 47 , and data not shown). However, it should be emphasized that, although no activation of in vitro PtdIns 5-P-producing activity was observed in response to insulin, PIKfyve’s functional significance to the insulin-induced elevation of PtdIns 5-P could be confined to relocation, as we have documented previously in 3T3-L1 adipocytes (48).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
The present studies identify PtdIns 5-P as a novel key signaling intermediate in acute insulin action on F-actin stress fiber disassembly and GLUT4 translocation in insulin-sensitive CHO-T cells and 3T3-L1 adipocytes, which operates by a PI 3-kinase-insensitive mechanism. This finding is based on several lines of experimental evidence: first, acute insulin markedly increased PtdIns 5-P levels in both CHO-T cells and 3T3-L1 adipocytes, which proceeded in a wortmannin-independent mechanism (Fig. 1Go); second, PtdIns 5-P, when elevated in CHO-T cells by PtdIns 5-P cytoplasmic microinjection or by PIKfyveWT ectopic expression, mimicked insulin’s effect on F-actin stress fiber breakdown (Figs. 3Go and 4Go); third, wortmannin pretreatment did not prevent the PtdIns 5-P-induced loss of F-actin stress fibers in these cells (Fig. 3Go); fourth, PtdIns 5-P was able to partially induce EGFP-GLUT4 translocation onto the 3T3-L1 adipocyte cell surface (Fig. 7Go); and fifth, both the insulin-induced loss of F-actin stress fibers and insulin-induced GLUT4 cell surface redistribution were largely abrogated in the presence of ectopically expressed or microinjected 3xPHD finger domain peptides that sequester functional PtdIns 5-P (Figs. 5Go, 6Go, and 8Go). Thus, PtdIns 5-P functions as a positive regulatory intermediate in insulin-regulated organization of F-actin and dynamics of GLUT4 vesicles.

A central role in actin filament remodeling stimulated by growth factors, such as insulin or IGF-1, has been attributed to the activated PI 3-kinase pathway and the subsequent activation of the small GTP-binding proteins (34, 49). However, more recent data indicate that insulin action relays two types of signals to modulate F-actin dynamics, one that is PI 3-kinase dependent and another that proceeds in a PI 3- kinase-independent manner. Thus, pharmacological inhibition of PI 3-kinase did not affect insulin action on F-actin stress fiber breakdown in CHO-T and 3T3-L1 fibroblasts, whereas insulin’s effect on membrane ruffling was abrogated (29, 30). Likewise, SHIP and GRP1, which hydrolyze or sequester the PI 3-kinase product PtdIns 3,4,5-P3, respectively, did not inhibit insulin’s effect on stress fiber disassembly in HIRcB cells, yet they both inhibited membrane ruffling (41, 42). Based on the data presented herein, we suggest that PtdIns 5-P selectively mediates the PI 3-kinase-independent insulin signaling to F-actin stress fiber breakdown but it is not involved in membrane ruffling. The latter requires activation of the PI 3-kinase pathway, as suggested previously (34, 50). The predicted role of PtdIns 5-P in the molecular mechanisms of F-actin stress fiber disassembly is further supported by recent studies demonstrating disappearance of actin stress fibers, along with membrane bleb formation, upon expression of the S. flexneri virulence factor IpgD in mammalian cells (17). However, IpgD expression is associated with rather complex changes in the overall PI intracellular metabolism, whereby, in addition to the rise of PtdIns 5-P and a massive PtdIns 4,5-P2 hydrolysis, an activation of the PI 3-kinase pathway occurs to increase higher 3'-PIs such as PtdIns 3,5-P2, PtdIns 3,4-P2, and PtdIns 3,4,5-P3 (17), each able to induce actin cytoskeleton remodeling on its own (8). The data presented herein, based on a direct and selective manipulation in PtdIns 5-P levels, provide compelling and unequivocal evidence to implicate PtdIns 5-P in the regulation of F-actin stress fiber dynamics. Moreover, loss of F-actin stress fibers was not produced by any of the microinjected 3'-PIs used here (Fig. 3Go, B and C), consistent with the notion that activation of the PI 3-kinase pathway does not play a role in this insulin effect. Rather, our data infer that an increase in PtdIns 5-P mass is both required and sufficient to induce the loss of actin stress fibers in response to insulin.

According to a currently held view, insulin-regulated translocation of GLUT4 vesicles from intracellular storage compartments to the 3T3-L1 adipocyte cell surface appears to require not only a PI 3-kinase-dependent but also a PI 3-kinase-independent cascade (22, 23, 24). Our observations for activated GLUT4 vesicle exocytosis to the cell surface in response to PtdIns 5-P microinjection in 3T3-L1 adipocytes (Fig. 7Go), together with insulin-induced increases in PtdIns 5-P in this cell type (Fig. 1Go), are consistent with the notion that the PtdIns 5-P pathway may be necessary in the insulin-activated PI 3-kinase-independent cascade. Studies in several laboratories have implicated the mobilization of actin filament rearrangement in insulin action on GLUT4 because various pharmacological agents that stabilize or disrupt F-actin structures are inhibitory (25, 26, 51). Recently it has been demonstrated that GLUT4 vesicle translocation in response to insulin action in 3T3-L1 adipocytes requires remodeling of two distinct compartmentalized F-actin populations (31). They both need activated TC10, a Rho family GTPase specific for 3T3-L1 adipocytes, and proceed in a PI 3-kinase-independent manner (31). Although we did not examine the effect of PtdIns 5-P on the complex F-actin reorganization in 3T3-L1 adipocytes in this study, it is conceivable that insulin-elevated PtdIns 5-P mass promotes an active F-actin breakdown as a first step to a complex F-actin remodeling for optimal efficiency of GLUT4 vesicle delivery to the adipocyte PM. Further work will be necessary to support a plausible relationship between the elevated PtdIns 5-P and activated TC10 pathway in response to insulin, as well as the relevance of a PtdIns 5-P pathway in insulin action on glucose uptake.

The enzymology underlying the observed robust insulin-dependent PtdIns 5-P increase was also addressed in this study, but due to the complex PI intracellular metabolism, the data still do not allow a definitive conclusion. We have previously shown that the synthetic arm in PtdIns 5-P production, at least in part, is due to PIKfyve, because ectopic expression of PIKfyveWT or the dominant-negative kinase-deficient PIKfyveK1831E mutant increases or decreases, respectively, the intracellular PtdIns 5-P levels in HEK293 stable cell lines (16). Consistent with these data, we have found here that PIKfyveWT expression in CHO-T cells partially reproduced the PtdIns 5-P effect in actin stress fiber breakdown (Fig. 4Go). However, the in vitro PIKfyve lipid kinase activity appears to be insensitive to insulin-directed activation in both 3T3-L1 adipocytes and CHO-T cells (Ref. 47 , and this study). Because acute insulin action in 3T3-L1 adipocytes causes a recruitment of cytosolic PIKfyve to membranes (48) where the PtdIns substrate resides, it is conceivable that elevation in PtdIns 5-P mass in response to insulin would be seen even in the absence of detectable increases in PIKfyve intrinsic activity. The role of the PIKfyve pathway in acute insulin action on glucose metabolism is further substantiated by data demonstrating inhibition of insulin-regulated GLUT4 vesicle translocation in 3T3-L1 adipocytes expressing dominant-negative kinase-deficient PIKfyveK1831E mutant (35). Thus, these data and considerations are consistent with the concept that the robust increase in PtdIns 5-P mass in response to insulin is due, at least in part, to PIKfyve.

Another source for a PtdIns 5-P mass increase is an augmented breakdown of PtdIns 4,5-P2. Such a pathway has been recently seen upon Shigella flexneri invasion of epithelial cells (17) and found to coincide with an active F-actin remodeling. Although a mechanism of PtdIns 4,5-P2 hydrolysis is plausible to operate in response to insulin, HPLC analyses of the PtdIns 4,5-P2 head group showed no significant decreases in PtdIns 4,5-P2 levels upon insulin stimulation of metabolically labeled 3T3-L1 adipocytes and CHO-T cells (Fig. 9Go). However, it is worth emphasizing that even subtle diminution of total PtdIns 4,5-P2 would produce a substantial PtdIns 5-P increase due to the high levels of intracellular PtdIns 4,5-P2 vs. PtdIns 5-P in these cell types (16, 37). Moreover, expected are spatially restricted changes, which may not result in an overall alteration in 32P-PtdIns 4,5-P2 accumulation. Furthermore, though currently unknown, the pathway of PtdIns 5-P hydrolysis could also represent a potential insulin-sensitive target to yield a PtdIns 5-P increase. Recent studies in Dictyostelium, identifying a new PtdIns 5-P-specific phospholipidinositol phosphatase PLIP (52), indicate that this assumption may be correct. Finally, the increase in PtdIns 5-P could be associated with insulin-dependent negative regulation of type II PIPK activities that consume PtdIns 5-P, converting it to PtdIns 4,5-P2. This hypothesis is supported by new data, published while this manuscript was in its final stages of preparation, demonstrating decreased insulin signaling to PI 3-kinase and Akt upon expression of type II PIPK in CHO cells (53). Clearly, whereas the enzymology of the rapid increase/attenuation of PtdIns 5-P is rather complex and may involve up- or down-regulation of more than one pathway, the data presented herein identify, for the first time, an insulin-regulated PtdIns 5-P pathway that signals to F-actin depolymerization and GLUT4 dynamics. PtdIns 5-P target proteins relevant to actin remodeling in the context of insulin action remain to be identified.


   Acknowledgments
 
We thank Linda McCraw for the excellent secretarial assistance. We thank Drs. Richard Anderson, Or Gozani, Junying Yuan, Jeff Pessin, Kostya Kandror, and Mike Czech for the kind gifts of His-Type IIß PIPK, EGFP-ING2-3xPHD, EGFP-ING2-3xPHD3K, EGFP-ACS-3xPHD, EGFP-GLUT4 or Myc-GLUT4 cDNAs, and GLUT4 antibodies.


   Footnotes
 
This work was supported by the National Institutes of Health (DK58058) and American Diabetes Association research grants (to A.S.). Part of this work was presented at the 63rd Scientific Session of American Diabetes Association, New Orleans, 2003.

Abbreviations: CHO, Chinese hamster ovary; EGFP, enhanced green fluorescent protein; F-actin, filamentous actin; FITC, fluorescein isothiocyanate; GroPIns, glycerophosphorylinositol; GST, glutathione-S-transferase; IM, intracellular membrane; -P, phosphate; -P2, bisphosphate; -P3, trisphosphate; PI, phosphoinositide; PIPK, PI phosphate kinase; PM, plasma membrane; PtdIns, phosphatidylinositol; PX, Phox Homology; TLC, thin-layer chromatography.

Received April 16, 2004.

Accepted for publication July 20, 2004.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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